Nuclear Starring for Confocal Microscopy

This protocol was contributed by Mark Running (University of California, Berkeley).

A useful way to observe the development of embryos, as well as meristems and young primordia developing at the shoot apex, is by confocal microscopy after staining the nuclei with propidium iodide (Clark et al. 1993; Running et al. 1995). The number of cells can be exactly quantified in a meristem or in young primordia. Because embryonic and meris-tematic cells are largely filled out by their nuclei, it is convenient to image only the nuclei.

This method allows analysis of whole-mount material, which is more easily reconstructed than sectioned material.

Materials

Tissue sample (embryos dissected from the seed coat or the growing tip of a plant)

Vacuum desiccator

In-house vacuum line or vacuum pump

Fixative (prepare fresh for each use)

15% (v/v) double-distilled water l-arginine buffer (0.1 m, pH 12.4 and pH 8.0) For pH 12.4: Adjust with 5 m NaOH. For pH 8.0: Adjust with HCl.

Propidium-iodide-staining solution (100 (ig/ml in 0.1 m l-arginine, pH 12.4)

Store wrapped in aluminum foil at 4°C (will last ~4-6 months). The solution is no longer usable when it turns a dull, dark orange color.

Decreasing and increasing ethanol series (95%, 85%, 70%, 50%, 30%, 15% and 15%, 30%, 50%, 70%, 85%, 95%, 100%)

High-quality ethanol (e.g., Gold Seal) or 100% bulk ethanol that has been dehydrated with molecular sieves (e.g., Sigma M 6141) (see Step 13).

Xylene series (75:25 ethanohxylene, 50:50, 25:75) and 100% xylene

Dissecting microscope

Equipment for fluorescence microscopy

CAUTION: ethanol, formaldehyde, HC1, propidium iodide, xylene (see Appendix 3)

Procedure

Fixation of Tissue

1. Place tissue in 10-15 ml of fixative in a glass scintillation vial.

2. Place the vials into a vacuum desiccator. Using an aspirator, apply full vacuum and then release slowly (15 minutes under vacuum is ideal).

3. Remove vials and check that all the tissue is submerged. If the tissue does not sink, reapply the vacuum.

4. Fix the tissue overnight (at least 4 hours) at room temperature.

5. Pour off fixing solution and replace with 70% ethanol.

A fine mesh screen helps to keep samples in the vial.

6. Subject the sample to a graded ethanol series (85%, 95%, 100%) for at least 30 minutes at each concentration.

7. Leave the sample overnight in 100% ethanol in a tightly capped vial.

This treatment will remove residual chlorophyll. Thicker tissue samples may require more time for complete chlorophyll elimination.

Staining

8. Subject the sample to a decreasing ethanol series (95%, 85%, 70%, 50%, 30%, 15%) for 30 minutes at each concentration. Transfer the sample to distilled water for a final 30-minute treatment.

9. Replace the distilled water with propidium-iodide-staining solution.

The volume of staining solution used is critical; use increasing volumes for larger quantities of tissue, ensuring that all parts of the sample are submerged. For apices, use -500 (il of staining solution per apex.

10. Stain the samples, out of direct light, for a minimum of 24 hours at 4°C.

For apices, a 4-day staining period works well, as long as the tissue has been fixed overnight. After the staining procedure, the tissue may appear slightly orange.

11. Rinse the sample with 0.1 m l-arginine buffer (pH 8.0).

12. Stand the sample in arginine buffer for 4 days at 4°C without agitation, changing the rinsing solution once every day.

Clearing

13. Subject the sample to a graded ethanol series (15%, 30%, 50%, 70%, 85%, 95%, 100%, 100%, 100%) for a minimum of 30 minutes at each concentration. Make the last two 100% ethanol changes with either a fresh bottle of high-quality ethanol or 100% bulk ethanol that has been dehydrated with molecular sieves.

14. Treat the sample with a xylene series (75:25 ethanohxylene, 50:50, 25:75) for a minimum of 2 hours at each concentration, followed by three changes of 100% xylene (at least 2 hours each).

Dissection of Apices

15. Place four individual drops of immersion oil on a microscope slide. The first drop should be large enough to contain the entire sample.

16. Use fine forceps to place the tissue sample , in the first drop and begin dissection under a dissecting microscope. For the shoot apex, hold the stem with one pair of forceps, and break off the older flowers by pushing them toward the base of the stem. Because the sample is dissected in solution, use strong indirect lighting. Direct overhead lighting, which results in glare, is unsuitable.

17. Dissect until the first drop of oil becomes too cluttered, then transfer the apex to the next drop of immersion oil for further dissection, increasing the magnification if necessary.

18. Continue dissection, transferring the material to a new drop of immersion oil as necessary, until dissection is complete and the sample is in the final drop of immersion oil.

For the study of the shoot apex, remove most of the visible floral primordia. This requires a magnification of 80x or more to be accomplished easily.

Mounting

19. After removing as much tissue as possible from the sample, use fine forceps to transfer it onto the center of a clean coverslip.

In many cases, multiple samples can be placed on the same coverslip.

20. Add a small amount of immersion oil to the sample by dipping forceps into a drop of oil, and transferring a droplet to the sample. Make sure that there is enough immersion oil to cover the sample, but not so much that it floats around.

21. For thicker tissues, adjust the position of the sample such that the area of interest is close to the surface of the coverslip. Shoot apices should abut the coverslip surface.

22. When the sample is correctly positioned, invert the coverslip and place it on a slide. For flat samples, such as mature leaves, sepals, or petals, use conventional microscope slides. For thicker samples, such as the shoot apex, use depression slides. In this case, place the coverslip obtained in Step 19 over the depression of the slide, with the sample attached to and below the coverslip but not touching the slide. Do not allow the immersion oil to touch the slide.

23. Seal the coverslip with nail polish. Viewing the Sample

Propidium iodide has a maximum absorption near 520 nm. Fluorescence can thus be excited by either a helium/neon laser, which emits at 543 nm, or an argon laser emitting at 514 nm. In most cases, the laser can be attenuated to 10% of full power; thus reducing the bleaching of the dye. Since the emission maximum of propidium iodide is at 610 nm, use a long-pass barrier filter to allow passage of light at wavelengths greater than 590 nm.

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